Name: murine cervical aortic transplantation model using a modified non-suture cuff technique
Uploaded: 2019-11-02T13:00:00
Description: Watch this Scientific Journal Video about Murine Cervical Aortic Transplantation Model using a Modified Non-Suture Cuff Technique at JoVE.com

All experiments were performed according to the guidelines of the German animal welfare act (TierSchG.) (AZ: 55.2-1-54-2532.Vet_02-80-2015).
1. Animal housing
<ol>
	<li>For experiments, use male C57BL/6 and BALB/c mice weighing 20-25 g with C57BL/6 mice as the recipient animals and BALB/c mice as the donor animals.</li>
	<li>Purchase the animals and house in a barrier pathogen-free facility, in accordance with the FELASA guidelines for health monitoring12.</li>
	<li>K...

<ol>
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R., Kaplan, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lack+of+improvement+in+renal+allograft+survival+despite+a+marked+decrease+in+acute+rejection+rates+over+the+most+recent+era.">Lack of improvement in renal allograft survival despite a marked decrease in acute rejection rates over the most recent era.</a> American Journal of Transplantation. 4 (3), 378-383 (2004).</li><li>Bagnasco, S. M., Kraus, E. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intimal+arteritis+in+renal+allografts:+new+takes+on+an+old+lesion.">Intimal arteritis in renal allografts: new takes on an old lesion.</a> Current Opinion in Organ Transplantation. 20 (3), 343-347 (2015).</li><li>Hollis, I. B., Reed, B. N., Moranville, M. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Medication+management+of+cardiac+allograft+vasculopathy+after+heart+transplantation.">Medication management of cardiac allograft vasculopathy after heart transplantation.</a> Pharmacotherapy. 35 (5), 489-501 (2015).</li><li>Verleden, G. M., Raghu, G., Meyer, K. C., Glanville, A. R., Corris, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+classification+system+for+chronic+lung+allograft+dysfunction.">A new classification system for chronic lung allograft dysfunction.</a> The Journal of Heart and Lung Transplantation. 33 (2), 127-133 (2014).</li><li>Ramzy, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiac+allograft+vasculopathy:+a+review.">Cardiac allograft vasculopathy: a review.</a> Canadian Journal of Surgery. 48 (4), 319-327 (2005).</li><li>Skoric, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiac+allograft+vasculopathy:+diagnosis,+therapy,+and+prognosis.">Cardiac allograft vasculopathy: diagnosis, therapy, and prognosis.</a> Croatian Medical Journal. 55 (6), 562-576 (2014).</li><li>Koulack, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+a+mouse+aortic+transplant+model+of+chronic+rejection.">Development of a mouse aortic transplant model of chronic rejection.</a> Microsurgery. 16 (2), 110-113 (1995).</li><li>Rowinska, Z., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Using+the+Sleeve+Technique+in+a+Mouse+Model+of+Aortic+Transplantation+-+An+Instructional+Video.">Using the Sleeve Technique in a Mouse Model of Aortic Transplantation - An Instructional Video.</a> Journal of Visualized Experiments. (128), (2017).</li><li>Dietrich, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mouse+model+of+transplant+arteriosclerosis:+role+of+intercellular+adhesion+molecule-1.">Mouse model of transplant arteriosclerosis: role of intercellular adhesion molecule-1.</a> Arteriosclerosis, Thrombosis, and Vascular Biology. 20 (2), 343-352 (2000).</li><li>M&#228;hler Convenor, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=FELASA+recommendations+for+the+health+monitoring+of+mouse,+rat,+hamster,+guinea+pig+and+rabbit+colonies+in+breeding+and+experimental+units.">FELASA recommendations for the health monitoring of mouse, rat, hamster, guinea pig and rabbit colonies in breeding and experimental units.</a> Laboratory Animals. 48 (3), 178-192 (2014).</li><li>Ollinger, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Blockade+of+p38+MAPK+inhibits+chronic+allograft+vasculopathy.">Blockade of p38 MAPK inhibits chronic allograft vasculopathy.</a> Transplantation. 85 (2), 293-297 (2008).</li><li>Thomas, M. N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=SDF-1/CXCR4/CXCR7+is+pivotal+for+vascular+smooth+muscle+cell+proliferation+and+chronic+allograft+vasculopathy.">SDF-1/CXCR4/CXCR7 is pivotal for vascular smooth muscle cell proliferation and chronic allograft vasculopathy.</a> Transplant International. 28 (12), 1426-1435 (2015).</li><li>Ollinger, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bilirubin:+a+natural+inhibitor+of+vascular+smooth+muscle+cell+proliferation.">Bilirubin: a natural inhibitor of vascular smooth muscle cell proliferation.</a> Circulation. 112 (7), 1030-1039 (2005).</li><li>Segura, A. M., Buja, L. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiac+allograft+vasculopathy:+a+complex+multifactorial+sequela+of+heart+transplantation.">Cardiac allograft vasculopathy: a complex multifactorial sequela of heart transplantation.</a> Texas Heart Institute Journal. 40 (4), 400-402 (2013).</li><li>McDaid, J., Scott, C. J., Kissenpfennig, A., Chen, H., Martins, P. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+utility+of+animal+models+in+developing+immunosuppressive+agents.">The utility of animal models in developing immunosuppressive agents.</a> European Journal of Pharmacology. 759, 295-302 (2015).</li><li>Shi, C., Russell, M. E., Bianchi, C., Newell, J. B., Haber, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Murine+model+of+accelerated+transplant+arteriosclerosis.">Murine model of accelerated transplant arteriosclerosis.</a> Circulation Research. 75 (2), 199-207 (1994).</li><li>Koulack, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Importance+of+minor+histocompatibility+antigens+in+the+development+of+allograft+arteriosclerosis.">Importance of minor histocompatibility antigens in the development of allograft arteriosclerosis.</a> Clinical Immunology and Immunopathology. 80 (3 Pt 1), 273-277 (1996).</li><li>Maglione, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+technique+for+heterotopic+vascularized+pancreas+transplantation+in+mice+to+assess+ischemia+reperfusion+injury+and+graft+pancreatitis.">A novel technique for heterotopic vascularized pancreas transplantation in mice to assess ischemia reperfusion injury and graft pancreatitis.</a> Surgery. 141 (5), 682-689 (2007).</li><li>Oberhuber, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Murine+cervical+heart+transplantation+model+using+a+modified+cuff+technique.">Murine cervical heart transplantation model using a modified cuff technique.</a> Journal of Visualized Experiments. (92), e50753 (2014).</li><li>Nakao, A., Ogino, Y., Tahara, K., Uchida, H., Kobayashi, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Orthotopic+intestinal+transplantation+using+the+cuff+method+in+rats:+a+histopathological+evaluation+of+the+anastomosis.">Orthotopic intestinal transplantation using the cuff method in rats: a histopathological evaluation of the anastomosis.</a> Microsurgery. 21 (1), 12-15 (2001).</li></ol>

Chronic allograft vasculopathy is the major cause of late graft loss after solid organ transplantation of the heart and likely renal and lung allografts8. Thus far, no causal therapeutic regimen could be developed in order to prevent the formation of CAV.
The pathophysiology of CAV is multifactorial and involves immunological and non-immunological aspects16. The use of rodent models in transplantation have been essential in understanding the unde...

Over the past six decades, solid organ transplantation has evolved from an experimental procedure to a standard of care for the treatment of end-stage organ failure1. Due to the improvement of antimicrobial agents, surgical techniques and advancement in immunosuppressive regiments, the early success rate of solid organ transplantation have significantly increased over the past decades2.
However, long-term graft survival rates have not significantly improved in the same manner3. The development of CAV is the major factor limiting long-term survival4,5,6. This pathology is characterized by the formation of a concentric neointimal layer consisting of smooth muscle cells, leading to progressive narrowing of the vessel and consecutive malperfusion of the transplanted solid organ. In heart transplant recipients, CAV lesions can be diagnosed in up to 75% of patients 3 years after transplantation7.
The pathophysiology of CAV is not fully understood yet. It seems to be related to numerous immunological and non-immunological factors, leading to endothelial damage with subsequent endothelial activation and dysfunction8. Thus far, no causal treatment option exists for the prevention of CAV, emphasizing the need for a reproducible small animal model in order to study the formation and potential therapy of CAV.
With the use of murine aortic transplantation models, CAV like lesions can be seen 4 weeks after transplantation. Those lesions consist mainly of vascular smooth muscle cells, thereby, resembling the human pathology. Because of a wide variety of transgenic and knock out mice, the use of mouse models in transplant associated pathologies offers a unique opportunity to identify new therapeutic options and understand their development. Due to the small diameter of the transplanted vessels however, the use of mouse models is commonly associated with long learning curves and an initial high complication rate9. With the introduction of the non-suture cuff technique, this most challenging part of the operation can be facilitated and the diameter of the anastomosis is kept constant10,11.

<table>
	<tbody>
		<tr>
			<td>Balb-c Mice (H2-d)</td>
			<td>Charles River</td>
			<td>Strain# 028</td>
			<td>Donor animal</td>
		</tr>
		<tr>
			<td>Bipolar cautery system</td>
			<td>ERBE</td>
			<td>ICC 50 / 20195-023</td>
			<td>Bipolar cautery</td>
		</tr>
		<tr>
			<td>C57BL/6J (H-2b)</td>
			<td>Charles River</td>
			<td>Strain# 027</td>
			<td>Recipient animal</td>
		</tr>
		<tr>
			<td>Halsey Needle Holders</td>
			<td>FST</td>
			<td>12501-12</td>
			<td>Needle Holder</td>
		</tr>
		<tr>
			<td>Halsted-Mosquito Forceps</td>
			<td>AESCULAP</td>
			<td>BH111R</td>
			<td>Curved Clamp</td>
		</tr>
		<tr>
			<td>Medical Polyimide Tubing</td>
			<td>Nordson MEDICAL</td>
			<td>141-0031</td>
			<td>Cuff-Material</td>
		</tr>
		<tr>
			<td>Micro Serrefines</td>
			<td>FST</td>
			<td>18055-04</td>
			<td>Micro Vessel Clip</td>
		</tr>
		<tr>
			<td>Micro-Adson Forceps (serrated)</td>
			<td>FST</td>
			<td>11018-12</td>
			<td>Standard Forceps</td>
		</tr>
		<tr>
			<td>Micro-Serrefine Clamp Applying Forceps</td>
			<td>FST</td>
			<td>18057-14</td>
			<td>Clipapplicator</td>
		</tr>
		<tr>
			<td>S&#38;T Forceps - SuperGrip Tips (Angled 45&#176;)</td>
			<td>S&#38;T</td>
			<td>00649-11</td>
			<td>Fine Forceps</td>
		</tr>
		<tr>
			<td>S&#38;T Vessel Dilating Forceps - Angled 10&#176; (Tip diameter 0.2 mm)</td>
			<td>S&#38;T</td>
			<td>00125-11</td>
			<td>Vesseldilatator</td>
		</tr>
		<tr>
			<td>Schott VisiLED Set</td>
			<td>Schott</td>
			<td>MC 1500 / S80-55</td>
			<td>Light</td>
		</tr>
		<tr>
			<td>Stereoscopic microscope</td>
			<td>ZEISS</td>
			<td>SteREO Discovery.V8</td>
			<td>Microscope</td>
		</tr>
		<tr>
			<td>Student Fine Scissors / Surgical Scissors - Sharp-Blunt</td>
			<td>FST</td>
			<td>91460-11 / 14001-12</td>
			<td>Standard Sissors</td>
		</tr>
		<tr>
			<td>Vannas-T&#252;bingen Spring Scissors (curved, 8.5 cm)</td>
			<td>FST</td>
			<td>15004-08</td>
			<td>Microsissors (curved)</td>
		</tr>
		<tr>
			<td>Vannas-T&#252;bingen Spring Scissors (straight, 8.5 cm)</td>
			<td>FST</td>
			<td>15003-08</td>
			<td>Microsissors (straight)</td>
		</tr>
	</tbody>
</table>

murine cervical aortic transplantation model using a modified non-suture cuff technique

With the introduction of powerful immunosuppressive protocols, distinct advances are possible in the prevention and therapy of acute rejection episodes. However, only minor improvement in the long-term results of transplanted solid organs could be observed over the past decades. In this context, chronic allograft vasculopathy (CAV) still represents the leading cause of late organ failure in cardiac, renal and pulmonary transplantation.
Thus far, the underlying pathogenesis of CAV development remains unclear, explaining why effective treatment strategies are presently missing and emphasizing a need for relevant experimental models in order to study the underlying pathophysiology leading to CAV formation. The following protocol describes a murine heterotopic cervical aortic transplantation model using a modified non-suture cuff technique. In this technique, a segment of the thoracic aorta is interpositioned in the right common carotid artery. With the use of the non-suture cuff technique, an easy to learn and reproducible model can be established, minimizing the possible heterogeneity of sutured vascular micro anastomoses.

In the fully MHC-mismatch transplantation model, a concentric neointimal layer can be seen 4 weeks after transplantation (Figure 2). This layer consists primarily of vascular smooth muscle cells as immunohistological staining for SM22 (a selective marker for mature vascular smooth muscle cells) revealed. As stated before, these vascular smooth muscle cells are pathognomonic for lesions seen in chronic allograft vasculopathy. For further analyses, aortic segments should be sectioned and stain...

Watch this Scientific Journal Video about Murine Cervical Aortic Transplantation Model using a Modified Non-Suture Cuff Technique at JoVE.com

Murine Cervical Aortic Transplantation Model using a Modified Non-Suture Cuff Technique

Here, we present a protocol of heterotopic aortic transplantation in mice using the non-suture cuff technique in a cervical murine model. This model can be used to study the underlying pathology of chronic allograft vasculopathy (CAV) and can help evaluate new therapeutic agents in order to prevent its formation.

With the introduction of powerful immunosuppressive protocols, distinct advances are possible in the prevention and therapy of acute rejection episodes. ...

I am Martin Ryll from the research laboratories of the Department of General, Visceral and Transplant Surgery of the LMU in Munich. Today we are going to show you a murine heterotopic aortic transplantation model using a modified non-suture cuff technique. First, anesthetize the recipient animal with an intraperitoneal injection of midazolam, medetomidine and fentanyl.Clip all hair of the cervical lateral region with an electric hair clipper for small animals and apply ophthalmic ointment with cotton swabs to prevent the eyes from drying out during the procedure. Place the animal in the supine position on a heating pad under the microscope and gently tape its legs to the operating table with skin sensitive plaster strips. Tilt its head back and scrub the operative field several times with alcohol.Make a skin incision from the jugular incision to the right lower mandible with small scissors. Remove the right lower lobe of the submandibular gland via bipolar cautery of the vessel pedicle and subsequent excision with micro scissors. Remove the right sternocleidomastoid muscle via bipolar cautery of the upper and lower portion and subsequent excision with micro scissors to gain access to the common carotid artery.Mobilize the common carotid artery as far distally and proximally as possible by pulling the surrounding connective tissue apart with fine forceps. Tie two 7-0 silk ligatures with minimal distance between each other around the middle of the common carotid artery and transect the common carotid artery with fine micro scissors between the ligatures. Pass the proximal ligated end through the cuff and fix it with a small artery clamp.Remove the ligature with fine micro scissors as close to the ligature as possible and flush the lumen with heparinized saline with a 30 gauge needle while taking care not to damage the vessel walls. Distend the open lumen using fine vascular dilatators and invert the carotid stump over the cuff by pulling it gently over the polyamide tube. Immediately fix the inverted carotid with a loosely pre-tied 7-0 silk loop.Perform the same procedure on the other end of the carotid artery with the other cuff. Pass the distal ligated end through the cuff and fix it with a small artery clamp. Remove the ligature with fine micro scissors as close to the ligature as possible and flush the lumen with heparinized saline with a 30 gauge needle while taking care not to damage the vessel walls.Distend the open lumen using fine vascular dilatators and invert the carotid stump over the cuff by pulling it gently over the polyamide tube. Immediately fix the inverted carotid with a loosely pre-tied 7-0 silk loop. Set the recipient animal aside and moisturize the upper two-third with saline until the aortic segment is explanted.Anesthetize the donor mouse in the same fashion as the recipient animal. Clip all hair of the abdominal and thoracic region with an electric hair clipper for small animals and apply ophthalmic ointment with cotton swabs to prevent the eyes from drying out during the procedure. Place the animal in the supine position on a heating pad under the microscope and gently tape its legs to the operating table with skin sensitive plaster strips.Scrub the operative field several times with alcohol. Perform a midline abdominal laparotomy with small scissors and push the intestines slightly upward to expose the inferior vena cava. Inject the IVC with one cc of heparinized saline using a 30 gauge needle.Cut the abdominal aorta and IVC below the renal arteries with small scissors to exsanguinate the donor animal. Loosely place a compress into the abdomen to absorb the blood. Perform a thoracotomy via bilateral diversion of the ribs with scissors and tilt the anterior chest wall cranially with a surgical clamp to expose the mediastinum.Cut the IVC and the esophagus directly above the diaphragm with micro scissors. Remove the heart and the lungs by titling them upward with forceps holding the cut IVC and esophagus and then excising them with micro scissors from the base to gain access to the thoracic aorta and the dorsal mediastinum. Mobilize the thoracic aorta from its surrounding tissue by pulling apart the surrounding connective tissue and fat with fine forceps while being careful not to damage any intercostal arteries.Cauterize all branches from the thoracic aorta with bipolar cautery forceps and excise the aortic segment between the diaphragm and the aortic arch using micro scissors. Flush the excised aortic segment with heparinized saline with a 30 gauge needle while taking care not to damage the vessel walls to remove any remaining blood or clots and transfer the graft to the recipient animal. Pull the proximal end of the donor aortic segment over the proximal cuff on top of the inverted carotid artery with fine forceps and immediately fix it with a loosely pre-tied 7-0 silk loop.Trim the distal free end of the aortic graft with micro scissors so that the graft length fits the distance between the two cuffs. Pull the distal end of the donor aortic segment over the distal cuff on top of the inverted carotid artery with fine forceps and immediately fix it with a loosely pre-tied 7-0 silk loop. Remove the distal clamp first to allow retrograde perfusion.After achieving hemostasis, remove the proximal clamp to complete the anastomosis. Finally, close the wound with 6-0 continuous suture. After 28 days, the animals are sacrificed and the grafts preserved for histological staining.As shown in these Van-Gieson's stainings of aortic sections, fully MHC mismatched allografted aortas from Balb/c donors to black six recipients developed a significant neointimal hyperplasia resulting in luminal narrowing. On the other hand, syngenic aortic transplants from black six donors to black six recipients developed only little neointimal hyperplasia. Immunofluorescence staining against SM22, a marker for vascular smooth muscle cells, as shown in this picture in green fluorescence, demonstrates that the neointima mainly consists of vascular smooth muscle cells.The murine aortic transplantation serves as a model for chronic transplantation vasculopathy which is the most common cause of late graft failure. In comparison to the orthotopic abdominal model, our modified cuff technique is less microsurgical demanding leading to a high success rate. In addition, due to the constant diameter of the cuff, variations of the vascular anastomosis as seen in other microsurgical transplant models can be neglected.Furthermore, the warm ischemic time is kept constant throughout the experimental groups as the recipient animal is prepared first before the aortic segment is explanted. Our modified model should help other laboratories to establish murine aortic transplantation.

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